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Phil. Trans. R. Soc. B (2011) 366, 3545–3553
doi:10.1098/rstb.2011.0073
Review
Quality control in the initiation of eukaryotic
DNA replication
John F. X. Diffley*
Cancer Research UK London Research Institute, Clare Hall Laboratories, South Mimms,
Herts EN6 3LD, UK
Origins of DNA replication must be regulated to ensure that the entire genome is replicated precisely once in each cell cycle. In human cells, this requires that tens of thousands of replication
origins are activated exactly once per cell cycle. Failure to do so can lead to cell death or genome
rearrangements such as those associated with cancer. Systems ensuring efficient initiation of replication, while also providing a robust block to re-initiation, play a crucial role in genome stability. In
this review, I will discuss some of the strategies used by cells to ensure once per cell cycle replication
and provide a quantitative framework to evaluate the relative importance and efficiency of individual
pathways involved in this regulation.
Keywords: DNA replication; cell cycle; S phase
1. INTRODUCTION
The genomes of eukaryotic cells are replicated from
many replication origins distributed along multiple
chromosomes during the synthetic or S phase of the
cell cycle. This parallel processing approach to tackling
DNA replication was significant in the evolution of
eukaryotes; it allows cells to replicate even very large
genomes in relatively short periods of time, and was,
therefore, crucial in supporting the increase in
genome size needed for complex multi-cellular life.
However, this required the solution to a serious bookkeeping problem: the cell must ensure that sufficient
numbers of origins are used in each S phase without
the re-use of any origin in a single cell cycle. The use
of an insufficient number of origins would leave
regions of the genome unreplicated before mitosis,
generating broken chromosomes and loss of genetic
information, while re-initiation from any origin
would lead to unbalanced increases in gene dosage
and long-lived, potentially fragile replication forks. In
this review, I will describe the basic mechanisms that
ensure once per cell cycle replication in eukaryotic
cells and explore some of the complexities in this regulation. I will also provide some quantitative arguments
to explain why this complexity exists. The reader is
referred to a number of excellent recent reviews for
further detail [1 – 8].
2. DESIGN OF THE SYSTEM
In bacterial and eukaryotic cells, a group of related
‘initiator’ proteins specifies where replication origins
will be located and then act to load hexameric DNA
*john.diffley@cancer.org.uk
One contribution of 16 to a Theme Issue ‘The cell cycle’.
helicases required to unwind DNA during DNA replication. There is a fundamental difference in the way
the helicases are loaded in these two systems and it is
this difference that is key to understanding the control
of eukaryotic replication [7]. In the Gram-negative
bacterium Escherichia coli, the dnaA protein acts as
the ‘initiator’ protein: it binds as an oligomeric filament to multiple specific sequences within the origin
of replication (oriC) and dictates where DNA unwinding, and hence replication, will begin [9,10]. dnaA is a
member of the AAAþ family of ATPases, and ATP
plays a crucial role in dnaA function [11]. Although
dnaA can bind to oriC without ATP, only the ATPbound form of dnaA can induce DNA melting in a
region adjacent to the dnaA-binding sites known as
the 13mers. dnaA, together with another AAAþ
protein dnaC, then loads one hexameric helicase
(dnaB) around each of the single strands of the
melted 13mers. Once the helicases are loaded, they
can begin to unwind DNA and bidirectional replisomes can be assembled on the unwound DNA.
In E. coli, the first step in replication, dnaA binding
and origin melting, is tightly regulated by a variety of
mechanisms that are critical for preventing the immediate re-initiation of replication (see [9,10] for further
discussion). Some mechanisms regulate the occupancy
of oriC by dnaA (e.g. SeqA, DatA) and some regulate
the nucleotide state of dnaA (RIDA, DARS,
b-clamp). Although a detailed description of replication
control in bacteria is beyond the scope of this review, a
few points are relevant and worth making. Firstly, as we
will see below, the use of multiple mechanisms to prevent re-replication is also a feature of eukaryotic DNA
replication. Secondly, many of the mechanisms involved
in preventing re-replication are not conserved between
different bacterial species. For example, in E. coli,
methylation of adenine residues in GATC sequences
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J. F. X. Diffley Review. DNA replication control
ORC
G2–M
CDK
Cdt1
Cdc6
ORC
G1
pre-RC
MCM MCM
DDK
CDK
P
P 11
3
P
45
2
G
P
ORC
G
P
Mcm10 45
7
G1–S
pol ε
45
RPC
pol α
cm
P
P
pol α
pol ε
10
4
4
10
cm
M
S
pol ε
M
G
pre-IC
G
45
Figure 1. Stepwise assembly of DNA replication complexes. The individual steps leading to the assembly of bidirectional replisomes is outlined. Names associated with each of the complexes are shown on the right: pre-RC, pre-replication complex; pre-IC,
pre-initiation complex; RPC, replisome progression complex. Cell cycle phases permissive for the individual steps are shown on
the left. For simplicity, some of the protein names have been abbreviated: 11, Dpb11; 3, Sld3; 7, Sld7; 2, Sld2; G, GINS;
45, Cdc45; 4, Ctf4. The shapes of many of the individual components are loosely based on three-dimensional reconstructions
from electron micrographs: ORC and Cdc6 are from Chen et al. [17], Mcm double hexamer is from Remus et al. [18],
Cdc45, Mcm2-7, GINS (CMG) are from Costa et al. [19], DNA polymerase epsilon (pol1) is from earlier studies [20,21],
and DNA polymerase a (pola) is from Klinge et al. [21]. The roles of CTF4 and Mcm10 in the RPC are inferred from earlier
studies [22,23]. Cyclin-dependent kinase (CDK) phosphorylations are shown in red, Dbf4-dependent kinase (DDK)
phosphorylations are shown in blue. The order of DDK and CDK in activating replication comes from earlier studies [24– 26].
by Dam methylase plays a critical role in preventing
re-replication [12,13]. Before replication, oriC is fully
methylated by Dam methylase, but immediately after
replication, because the nascent DNA strand is
unmethylated, the double-stranded origin is transiently
hemi-methylated before Dam methylase can methylate
the nascent DNA. The SeqA protein binds tightly to
this hemi-methylated DNA, preventing dnaA from
re-binding to its weaker binding sites, thus preventing
re-initiation of replication [13–15]. Although the
Dam/seqA system is clearly important in E. coli and
mutants lacking this system inappropriately re-initiate
replication, this system is entirely absent from other
bacterial groups such as the Gram-positive bacteria
including Bacillus subtilis. Again, as we will see below,
the apparently rapid evolution of re-replication control
is also a feature of eukaryotic replication.
Phil. Trans. R. Soc. B (2011)
In eukaryotes, the initiator protein is a multisubunit protein called the origin recognition complex
(ORC; figure 1) [16]. Five of the six ORC subunits
are members of the AAAþ family (though only one,
Orc1, has retained a functional ATPase [27]) in a
clade of initiator proteins that includes dnaA [28].
Analogous to cooperation of dnaA with the AAAþ
dnaC, ORC cooperates with another AAAþ ATPase,
Cdc6, to load the replicative helicase, Mcm2-7, onto
origin DNA [29,30]. An additional factor, Cdt1,
which does not have an obvious bacterial analogue,
is also essential for helicase loading [31 – 33]. ATP
also plays a crucial role in ORC function; however,
this role is quite different from the role of ATP in
dnaA function. ATP binding (but not hydrolysis) is
required for budding yeast ORC to bind to its cognate
sequence within origins; ADP cannot fulfil this
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Review. DNA replication control J. F. X. Diffley 3547
requirement [16]. In contrast to dnaA, the binding of
ORC to origin DNA does not induce measurable
origin melting in the region of helicase loading.
Whereas ATP hydrolysis by dnaA primarily plays a
regulatory role by preventing immediate re-initiation,
ATP hydrolysis by ORC and Cdc6 is required for
the loading of the hexameric Mcm2-7 helicase onto
DNA [34,35].
The most important difference with bacteria derives
from this fact that ORC does not melt DNA. Instead,
it loads Mcm2-7 as a head-to-head double hexamer
around double-stranded DNA, not single-stranded
DNA [18,36]. In this configuration, in contrast to
the situation in bacteria, the helicase is inactive.
Indeed, this pre-replicative complex (pre-RC) can
reside at origins for significant periods of time before
the helicase is activated and DNA replication commences [37]. Activation of the helicase is a complex
and still poorly characterized step that involves the
reconfiguration of Mcm2-7 so that the double hexamer
separates and the individual hexamers are bound
around each of the single strands of DNA [2,38,39].
This step requires the loading of two additional proteins,
Cdc45 and GINS, onto Mcm2-7 in a reaction requiring
another set of factors including Sld2, Sld3, Sld7 and
Dpb11 [2]. The Cdc45–Mcm2-7–GINS complex
then acts as the replicative helicase [40], where it is
essential for DNA unwinding during the elongation
phase of DNA replication [41– 46].
The division of the initiation reaction into two
discrete steps (helicase loading and helicase activation) in eukaryotes has profound implications for the
control of DNA replication [7]. By differentially regulating these two steps so that they are restricted to
different phases of the cell cycle, cells can ‘license’
hundreds or thousands of origins with the inactive
helicase, and these origins can subsequently be
‘fired’ by helicase activation. Because the helicase
cannot be reloaded in S phase, origin firing is limited
to once per cell cycle. In general, licensing can occur
from the end of mitosis until a point in late G1
phase, and the helicase activation step can only occur
during S phase and G2 phase. The mechanisms by
which these steps are regulated during the cell cycle
are considered in §3.
3. REGULATION OF LICENSING
Pre-RC assembly is restricted to late M/early G1 phase
by at least three separate systems in eukaryotes.
In some organisms, all three are used, whereas in
others only one or two. These systems include
geminin-dependent inhibition of Cdt1 function,
Cul4 (Crl4)- and proliferating cell nuclear antigen
(PCNA)-dependent degradation of Cdt1 during S
phase and cyclin-dependent kinase (CDK)-dependent
inhibition of licensing via a variety of targets. These
have been the subject of a number of very thorough
reviews and the reader is encouraged to consult these
for more detail [1 – 4,6 – 8,47 – 49].
Geminin is a small, coiled-coil protein originally
identified in Xenopus egg extracts as a substrate for
the anaphase-promoting complex/cyclosome (APC/C)
[50] and an inhibitor of Cdt1 function in licensing
Phil. Trans. R. Soc. B (2011)
[51,52]. Because the APC/C is specifically active
during mitosis and G1 phase, geminin is inactivated
during this period, allowing Cdt1 to participate in
the licensing reaction only during this period. Geminin
appears to be found only in metazoans, where it contributes to preventing re-replication. Depletion or
deletion of geminin induces a G2/M checkpoint in
many different cell types. In some cell types, this is
accompanied by substantial re-replication, whereas in
other cell types it is accompanied by S phase delays.
Although these phenotypes appear superficially contradictory, they probably derive from the same root
cause: induction of re-replication. In some cases,
large numbers of origins are deregulated, resulting in
significant amounts of re-replication. In others, only
a few origins may be deregulated resulting in very
small amounts of re-replication [53]. In some cases,
this may even be manifested as an apparent reduction
in overall amounts of DNA replication [53] presumably
because checkpoint activation caused by re-replication
can prevent all new initiation, leading to an overall
shut down of replication. In general, cancer cells
appear especially prone to re-replication after geminin
depletion [54,55].
The second system contributing to preventing
re-replication involves the targeting of Cdt1 for degradation during S phase by an E3 ubiquitin ligase
containing Cul4 (Crl4), Ddb1 and Rbx1 and using
the Cdt2 substrate recognition subunit [56 – 58]. In
this system, Cdt1 is recruited to chromatin specifically
during S phase by interaction with the PCNA sliding
clamp processivity factor where it is ubiquitylated
and destroyed [59,60]. This system elegantly couples
the prevention of re-replication directly to the act
of replication and, as a consequence, operates only
during S phase of a normal cell cycle. Cul4-dependent
Cdt1 degradation has been conserved from fission
yeast through metazoans [61,62].
The final system working in most eukaryotes is
the only system operative in the budding yeast
Saccharomyces cerevisiae. This system involves direct
inhibition of pre-RC components by CDKs (reviewed
in [63]). Because CDKs are inactivated at the end of
mitosis and become re-activated in the late G1 phase,
this establishes a window of time during G1 phase
when licensing can occur. In budding yeast, where
this has been best characterized, Cdc6, ORC and
Mcm2-7 are all directly inhibited by CDK phosphorylation, each by different mechanisms. The
Cdc6 protein is phosphorylated by both the G1
phase cyclin (CLN)-associated CDK as well as the
S/G2/M phase cyclin (CLB)-associated CDK [64 –
67]. CDK phosphorylation of Cdc6 generates two
distinct binding sites for the Cdc4 subunit of the
Skp, Cullin, F-box containing complex (SCF) ubiquitin ligase [66]. Thus, CDK phosphorylation of Cdc6
inhibits its function by targeting it for ubiquitinmediated proteolysis. Later in the cell cycle, the mitotic
cyclin Clb2 binds tightly to CDK-phosphorylated Cdc6
and prevents it from associating with ORC [68]. The
Clb2-binding site overlaps one of the SCF-binding
sites, and stabilizes the Cdc6 protein in an inactive
form [66,68]. The degradation of Clb2 at the very end
of mitosis by the APC/C releases Cdc6 from its inhibitory
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J. F. X. Diffley Review. DNA replication control
complex and allows licensing to occur. CDKs inhibit
the Cdt1/Mcm2-7 complex by promoting its export
from the nucleus [69,70]. This, again, is accomplished
by both CLN- and CLB-associated CDK [70] and
involves direct phosphorylation of Mcm subunits [71].
Finally, ORC is phosphorylated on two subunits (Orc2
and Orc6) specifically by the CLB-associated CDK
[72,73]. ORC phosphorylation inhibits pre-RC assembly
by interfering with the interaction between ORC and
Cdt1 [74].
In addition to this role in inhibiting pre-RC assembly, CDKs play a second essential role in regulating
DNA replication: they are required to trigger initiation
from licensed origins. They do this in budding yeast by
phosphorylating Sld2 and Sld3 [75 – 77]. Phosphorylation of these proteins generates binding sites for
tandem BRCT repeats in the Dpb11 protein. Recently,
it has been shown that essential CDK phosphorylation
has been conserved in the human homologue of Sld3,
Treslin/ticrr [78,79]. As a consequence of these two
distinct roles for CDKs, pre-RC assembly is restricted
to G1 phase, when origins cannot fire because CDKs are
absent, and activation of CDK at the end of G1 phase
triggers initiation from licensed origins, and prevents
the re-assembly of pre-RCs at origins that have fired.
The role for CDK in preventing licensing outside of
G1 phase has been conserved in evolution; however,
the specific details of how CDK inhibits licensing
are quite different in different organisms. Chemical inhibition of CDKs in G2 and mitosis or genetic depletion of
the mitotic CDK1 promote re-licensing and allow
additional rounds of replication in human tissue culture
cells [80–82], very similar to re-replication induced
by deletion of mitotic cyclins in fission yeast [83] and
transient inhibition of CDK by overexpression of CDK
inhibitors in both fission and budding yeasts [84,85].
Moreover, in at least some cell types, cyclin A depletion
increases the amount of re-replication caused by geminin depletion [86]. Some targets of CDK inhibition
in human cells have been identified: both Orc1 and
Cdt1 can be targeted for SCFSkp2-dependent degradation [59,87] and Cdc6 phosphorylation can cause
its export from the nucleus [88–91]. Because of the
importance of geminin and Crl4-dependent Cdt1
degradation and because these pathways do not exist
in budding yeast, regulation of pre-RCs by CDKs in
metazoans has been somewhat understudied, and
further work is required to understand its importance
relative to the other two pathways.
Given the importance of preventing re-initiation of
DNA replication within a single cell cycle, it might
seem odd that details of this critical mechanism have
not been conserved in evolution. It is likely that two
factors contribute to the rapid evolution of licensing
regulation by CDKs. The first is the high level of overlap
built into the system. For example, while Cdc6 is well
established as a CDK target in budding yeast, mutation
of any individual phosphorylation site in Cdc6 does not
induce re-replication or result in significant reduction in
fitness. Indeed, a version of Cdc6 in which all CDK sites
have been eliminated is viable, grows apparently normally and does not show defects in origin function
as indicated by plasmid maintenance [64]. Similarly,
forced nuclear localization of Mcm2-7 throughout the
Phil. Trans. R. Soc. B (2011)
cell cycle or expression of unphosphorylatable orc
mutants alone does not induce detectable re-replication
even using sensitive comparative genome hybridization
methods [92]. It is only when deregulated components
are combined that detectable re-replication occurs. For
example, expression of stabilized Cdc6 together with
unphosphorylatable ORC is lethal and induces reinitiation from a subset of replication origins [68,72,
92]. It is only when all three proteins are deregulated
that substantial amounts of DNA re-replication can be
detected, for example, by flow cytometry [73].
The second factor that contributes to rapid evolution
is the interchangeability of regulatory mechanisms [93].
For example, although combination of stable Cdc6 with
an unphosphorylatable ORC is lethal, this lethality is
suppressed by fusion of a cell cycle-dependent degron
onto the Cdt1 protein, which confers CDK-dependent
degradation of Cdt1 during S, G2 and M phases
[93]. Also, addition of a cassette that confers CDKdependent nuclear export onto stable Cdc6 is sufficient
to restore viability when combined with unphosphorylatable ORC [93]. Thus, it appears that the molecular
mechanisms by which each pre-RC component is
inhibited by CDK are relatively unimportant; what is
important is that multiple pre-RC components are
inhibited by different mechanisms.
4. THE QUALITY CONTROL PROBLEM
To understand why so many mechanisms are involved
in preventing re-initiation, it is useful to consider the
scale of the problem: in cells with large genomes,
such as humans, re-initiation needs to be prevented
at tens of thousands of replication origins in each cell
cycle over the course of billions of cell cycles. Thus,
the block to re-replication needs to be extraordinarily
efficient. In the following section, I will examine the
implications this scale has on the problem. This was discussed in further detail in a previous review [94]. I will
initially consider the issue in budding yeast, making a
few simple assumptions. Firstly, DNA replication in
yeast initiates from approximately 400 origins during
each S phase, and re-initiation from any of these origins
counts as re-initiation. In human cells, the number is
approximately 50 000. Secondly, although the probability of re-initiating DNA replication is a function of
both the probability of re-licensing origins and the
probability of firing these re-licensed origins, for simplicity, we will set this second probability to ‘1’ (i.e. any
origin inappropriately re-licensed will re-initiate). Therefore, re-replication is entirely a function of inappropriate
re-licensing. Thirdly, each origin acts independently.
That is, each origin has some probability of re-initiating
that is unaffected by events at other origins.
With these assumptions, the probability that any
individual origin will re-initiate in a single cell cycle
can be converted into a probability that at least one
origin in the genome will re-initiate as follows: if p is
the probability an individual origin will re-initiate in
one cell cycle, q is the probability an individual origin
will not re-fire in one cell cycle and n is the number of
origins (400 in yeast), then:
ð p þ qÞn ¼ 1;
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Review. DNA replication control J. F. X. Diffley 3549
expanding this yields:
n
n
‘pindividual’, then:
p þ þ q ¼ 1;
pall ¼ ð pindividual Þ3
where p n is the probability that all origins will re-fire
while q n is the probability that no origin will re-fire in
a single cell cycle. All intermediate terms (not shown)
are the probabilities of different numbers of origins
re-firing. Thus, if the probability that a single origin
will not re-fire in a cell cycle is 99 per cent (0.99),
that is, the block to re-initiation per origin is 99 per
cent efficient in a single cell cycle, then the probability
that no origin will re-fire in yeast is (0.99)400 ¼ 0.018,
or approximately 2 per cent. In human cells, this is
(0.99)50 000 ¼ 6 102219! Thus, to achieve a robust
block to re-replication in each cell cycle, the block to
re-initiation on a per origin basis must be far, far greater
than 99 per cent. To achieve a 99 per cent probability that
no origin will re-fire in a single cell p
cycle
ffiffiffiffiffiffiffiffiffiffiin yeast, the
probability on a per origin basis is 400 0:99 ¼ 0:99998,
or 99.998
per
pffiffiffiffiffiffiffiffiffi
ffi cent efficient. In human cells,
this is 50 000 0:99 ¼ 0:9999998. Or, in other words, an
error rate of approximately 1 in 105 initiation events in
yeast and approximately 1 in 107 in human cells is
required to achieve this 99 per cent probability. Given
the fact that even very limited re-replication is lethal in
yeast [68,72,92], it is highly likely that the overall block
to re-initiation in wild-type cells is considerably greater
than 99 per cent.
or
5. A POSSIBLE SOLUTION
From the preceding discussion, it is clear that the
effective error rate in preventing re-initiation is likely
to be even lower than 1027 per origin. This approaches
the kinds of error rates observed in nucleotide insertion during DNA replication, and it is worth
comparing the systems. In the case of nucleotide
incorporation, accuracy is achieved by a series of
sequentially acting biochemical quality control mechanisms: replicative DNA polymerases have very high
levels of accuracy in initial incorporation, they also have
additional ‘proofreading’ exonucleases that can remove
misincorporated bases immediately after insertion, and
mismatch repair can catch any misincorporation that
slips through these first two mechanisms [95]. In the prevention of re-initiation, mechanisms do not appear to act
sequentially, but rather act in parallel. Nonetheless, the
outcome is similar: extraordinary accuracy.
To understand how parallel mechanisms cooperate
quantitatively, we assume that each mechanism operates independently of the other mechanisms. Using
budding yeast as an example, this means that Cdc6
degradation is independent of Mcm2-7 nuclear
export, which, in turn, is independent of ORC phosphorylation, etc. Importantly, re-initiation will only
occur at an origin if all mechanisms fail. So, if we
consider three separate mechanisms, each with a
probability of failing (pCdc6, pORC, pMcm), then the
probability that all three will fail (pall) is:
pall ¼ pCdc6 pORC pMcm :
For simplicity, if we assume that all mechanisms operate with similar efficiency, so pCdc6 ¼ pORC ¼ pMcm ¼
Phil. Trans. R. Soc. B (2011)
pindividual ¼
ffiffiffi
p
3
pall
so, to achieve an overall error rate per origin of 1025
needed in the budding yeast example above, then each
pathway needs an error rate of 0.02. In other words,
each pathway needs to be ‘only’ 98 per cent efficient.
The multiplicative relationship described above for
individual mechanisms blocking re-replication suggests
an interesting relationship between genome size and
mechanisms preventing re-replication: addition of just
one mechanism preventing re-initiation operating at
approximately 99 per cent efficiency is required for
every 100-fold increase in the number of origins used.
Assuming origin spacing is similar, this means an
additional mechanism can afford an organism a
100-fold increase in genome size.
6. PERSPECTIVES AND CHALLENGES
The calculations described above are, by necessity,
not based on any ‘real’ numbers. For example, what is
the actual rate of re-initiation in wild-type cells in vivo?
Previous plasmid loss assays in yeast have suggested
that the rate of re-initiation per origin is less than 1 in
1023 [96], which was the limit of detection in this
assay. However, as described above, the real number is
likely to be much lower than this. Similarly, just how efficient is the ubiquitin-mediated degradation of Cdc6 in
vivo? Or, how efficient is the export of Mcm2-7? Are
they 90, 99, 99.99 per cent efficient? Assays that can
measure intracellular concentrations of proteins over
the ranges required simply do not exist. Hopefully, the
development of more sensitive, quantitative assays for
these parameters will be available in the future to allow
re-examination of the issues described in this review.
Finally, how important is any of this? Work in yeast has
shown that mutations that decrease replication-initiation
efficiency lead to greatly elevated rates of gross chromosome rearrangements [97]. Moreover, mutations that
induce even small amounts of re-initiation lead to cell
death [68,72] or elevated rates of gene amplification
[98]. As a consequence, deregulated licensing has the
potential to drive genome instability. Indeed, overexpression of Cdt1 and Cdc6 in mouse models has been
shown to induce tumours [99,100]. Deregulated CDK
expression is common in cancer and has been shown to
inhibit licensing in both yeast and human cells [101–
103]. Thus, the ability to initiate replication efficiently
and, at the same time, efficiently prevent any re-initiation
may be a critical barrier to the development of cancer.
I am grateful to members of my laboratory for discussion.
Work in my laboratory was funded by Cancer Research
UK and the European Research Council grant 249883—
EUKDNAREP.
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